The Outer Banks are a group of barrier islands distributed in a north–south 320 km chain along the northeastern Coast of NC that separate the Albemarle, Pamlico and Currituck sounds from the Atlantic Ocean (Inman and Dolan, 1989). Questing ticks were initially collected at six sites in October 1991 (Fig. 1) (GPS coordinates are provided below). Additional adult tick collections were made at Audubon Pine Island Sanctuary (Pine Island) during 1992–1993 from September through March, encompassing the yearly period of activity of adult questing I. scapularis. Questing ticks were collected again in 2009, from four sites previously sampled between 1991 and 1993. Rodents were trapped and examined for ticks and spirochaetes at Pine Island and Buxton Woods between 1992–1994 and 2005.

The sites spanned the full length of the Outer banks in North Carolina along the main connecting highway, NC 12 (Fig. 1). Soils were consistently sandy, supporting predominately xeric vegetation typical of barrier islands that receive salt spray from the Atlantic, with two exceptions noted below at Pine Island and Nags Head Woods Ecological Preserve (Nags Head Woods). Marked within-site variation in plant species were apparent; example representative species and approximate site coordinates for collection sites are provided below. Corolla (36°22′52.00″N, 75°50′00.00″W), the most northern site is a small residential community with sandy soils and predominately mixed deciduous and pinewoods. Pine Island (36°13′42.14″N, 75°46′26.06″W) is an Audubon wildlife sanctuary located on the western side of highway NC-12 adjacent to Currituck Sound and the Intracoastal Waterway. The sanctuary is separated by a wide pedestrian greenway/road that is periodically mowed. Vegetation on the western side of the road is predominately maritime wet grassland (Schafale and Weakley, 1990). The eastern side of the sanctuary is predominately covered by a xeric maritime shrub thicket or salt shrub (Schafale and Weakley, 1990) that is dominated by bayberry (Myrica pensylvanica), live oak (Quercus virginiana) and wax myrtle (Morella cerifera) interspersed with loblolly pine (Pinus taeda), cedar (Juniperus virginiana) and mixed beach grasses (e.g. Ammophilia breviligulata, Uniola paniculata). Nags Head Woods (35°59′28.06″N, 75°39′46.95″W), which is protected by the relatively high dunes of Jockey’s Ridge State Park, a maritime deciduous forest of approximately 1000 acres in size with a diverse mixed flora of about 300 plant species. The forest supports 84 species of trees and shrubs, including loblolly pine, hickory (Carya spp.) oak (Querus spp), cedar (J. virginiana) and magnolia (Mangolia grandiflora). The understory includes 11 fern species (e.g. Amphicarpaea bracteata), goldenrod (Solidago canadensis), milkweed (Asclepias spp.), 45 species of vines and numerous other plant species (Krings, 2010). Ticks were also collected south of Nags Head Woods, adjacent to Coquina Beach (35°49′55.56″N, 75°33′41.82″W and at Bodie Island (35°53′39.62″N, 75°35′22.61″W), a connected peninsula covered by similar xeric salt shrub thicket and beach grasses. In Buxton Woods (35°15′41.16″N, 75°32′19.56″W) located at approximately 56 miles south of Nags Head Woods, sampling for questing adults was conducted in vacant lots with mixed grasses that were dispersed between residences. The sandy soils supported mixed grasses (e.g. Panicum amarum) and xeric scrub brush (e.g. Helianthemum canadense) (Stalter and Lamont, 1997). Sampling in Cape Hatteras was conducted at sites in Cape Hatteras National Seashore National Park (35°41′03.55″N, 75°18′04.43″W) in salt scrub thickets comprised of wax myrtle (M. cerifera), eastern baccharis (Baccharis halimifolia) and lanceleaf greenbrier (Smilax smallii). A single I. scapularis adult was collected at Pea Island National Wildlife refuge (35°41′30″N 75°32′45″W), a site with a vegetation shrub thicket and beach grasses. Sampling at the site was abruptly curtailed due to severe biting fly strike.

Adult questing I. scapularis were collected from vegetation with corduroy cloth. Approximately 1 × 1.5 m of corduroy suspended from a wooden base was dragged across the top of and through vegetation while walking. Actual dragging/flagging technique varied with the collector and the density and type of vegetation. On Pine Island, collections were predominately made along pedestrian hiking paths by dragging the cloth across the edge vegetation bordering the main access road and probing with the cloth along deer and rabbit trails. Ticks collected for culture or indirect immunofluorescence microscopy (IFA) were placed in a glass vial, containing a moistened Plaster of Paris/charcoal base, covered with a Nitex^® (Houston, TX, USA) mesh screen. The vial was placed in Whirl-Pak^® plastic pouch (Nasco, Fort Atkinson, WI, USA) with a lightly moistened cotton ball and then transferred to a cooler, containing reusable cooler packs or held at ambient temperature for transport to the laboratory. In the laboratory, ticks retained for culture or IFA testing were held in humidified plastic chambers below ambient temperature at approximately 20–21°C. Ticks collected for PCR testing were placed in 95% ethanol. Collected ticks were morphologically identified to species using standard dichotomous keys (Cooley and Kohls, 1945; Sonenshine, 1979).

A representative sample of adult ticks (n = 28), submitted for species validation to James Keirans (Georgia Southern University, Statesboro, GA), was confirmed to be I. scapularis and retained in the US National Tick collection as voucher specimens (Accession Numbers: 46104-10, 46116-46119, 46121-27).

All rodent collections were made at Pine Island and Buxton Woods. In all studies, rodents were trapped in Sherman live traps placed in lines approximately 7 m apart or in 7 × 7 grids. Traps were baited with a mixture of peanut butter and dry rolled oats, and cotton was included in the traps during colder months for insulation (Adler and Wilson, 1983). Trap locations were marked with flagging to aid in trap recovery.

Traps were set before dusk in the evening and then checked for rodents the next morning. Rodent trapping during January 1992–September 1993 on Pine Island was conducted with 49–100 Sherman live traps for four consecutive nights (2546 trap-nights). Additional trapping sessions were conducted for 1–3 consecutive nights with 50–57 traps in 1993–1994 (1461 trap-nights), and 2002 (799 trap-nights). Rodent trapping was conducted again in March–May 2005 (350 trap-nights). In these studies, 100 Sherman live traps were set the first night. Cold weather resulting in some rodent mortality prompted early curtailment of trapping after one night in March. High trapping success also prompted a reduction in the number of traps/night to 60 traps, for two nights, to accommodate handling by a single investigator.

Rabbits were trapped by placing two-door funnel wooden box traps or Havahart^® traps (Lititz, PA, USA) in lines in tall vegetation in readily distinguishable rabbit runs. No bait was used. Traps were set in the evening, checked in the morning and kept closed throughout the day.

Lizards were trapped using coverboards (0.6 m × 1.2 m × 1 cm exterior grade plywood) (Grant et al., 1992) and funnel traps placed at the end of drift fences (50 m length × 30.5 cm high) (Gibbons and Semlitsch, 1981). Drift fences were deployed in three lines in an H-shaped pattern. Funnel traps were opened in the morning, checked 2–3 times during the day and closed at night. Coverboards were placed in 7 × 7 grids (n = 49 per grid) and checked 2–3 times each day.

Restraint, handling, euthanasia and sampling protocols of trapped animals varied during the 18-year study period based on the samples to be collected, and changes in recommended anaesthetic protocols for rodents (IACUC-05-038-0). During studies conducted in 1991 and the spring of 1993, trapped animals were released into a bucket and restrained. Ticks were removed and placed in glass mesh-covered vials (noted above). A blood sample was obtained from the tail vein of each animal, after which the animals were identified with numbered ear tags, and released at the exact site of capture.

Captured lizards were identified and marked by toe-clipping (Ferner, 1979). Blood samples were placed in coolers containing reusable cold packs for transport to the laboratory and refrigerated upon arrival.

During August 1993, and monthly during October 1993 through September 1994, trapped rodents were transported to the laboratory for tick collection, euthanasia and blood culture. Rodents were transferred to stainless cages with a mesh bottom suspended over water to collect ticks dropping off of hosts. The ticks were collected from the water and identified to species. After the ticks had dropped off their hosts, the rodents were euthanized with carbon dioxide. Samples of cardiac blood, ear skin and bladder were aseptically obtained and inoculated into 8-ml tubes of BSK-II media and antibiotics (8 mg/ml kanamycin sulphate and 230 mg/ml of 5-fluoruracil (Barbour, 1984). Embryos were obtained from two white-footed mice and one rice rat during necropsy. Embryonated tissues were inoculated into BSK-II after rinsing in 70% alcohol and sterile PBS and dissecting in sterile petri dishes.

Additional rodent trapping was conducted during May and July 2002. Animals were anesthetized via intraperitoneal injection of ketamine (100 mg/kg) and xylazine (10 mg/kg) mixture and euthanized via cervical dislocation in accordance with the AVMA Guidelines for the Euthanasia of Animals (AVMA 2001). Blood (obtained by cardiac puncture) and bladder tissue were collected for spirochaete culture.

During March–May 2005, trapped rodents were anesthetized with midazolam (5 mg/kg), and medetomidine (0.5–1 mg/kg) via intraperitoneal injection. After sampling, the rodents were administered Flumazenil (0.5 mg/kg) and Atipamezole (2 mg/kg) intramuscularly for reversal of sedation and then released after recovery. Trapped rabbits were anesthetized using a combination of midazolam 0.5 mg/kg and butorphanol 0.2 mg/kg. Once sedated, a 1 ml blood sample was obtained from the jugular vein for serum. Atipamezole 1 mg/kg intramuscularly was given to reverse the anaesthesia, and rabbits were released in the same area that they were trapped after full recovery from anaesthesia. Ear punch biopsies (Sinsky and Piesman, 1989) were obtained to attempt culture of B. burgdorferi. The pinnae were cleaned with 95% ethanol, and an ear punch biopsy was obtained from one pinna using a 1-mm stainless steel biopsy tool. The specimen was then rinsed in sterile phosphate buffered saline (pH 7.38) (Sinsky and Piesman, 1989). The tissue was minced with a pair of sterile scissors and then placed in BSK-II medium for culture as described below.

Protocols used to culture B. burgdorferi relied on variations of BSK-II media (Barbour, 1984). During 1991–1993, BSK-II was used for attempted culture using procedures originally developed by Barbour (1984) and modified by Steere et al. (1983). During 2002 and 2005, BSK-H (Pollack et al., 1993) was used for attempted B. burgdorferi culture. During 1991–1994, we attempted to culture spirochaetes from questing ticks and hosts. Prior to dissection, ticks were cleansed and disinfected by passing them through a series of baths: Betadine solution [Purdue Pharma, Stamford, CT], PBS, hydrogen peroxide, PBS, 70% alcohol and PBS. Culture tubes were held at 35°C and examined for spirochaete growth by darkfield microscopy at 3 days post-culture and then at 1 week, and then consecutively at weekly intervals thereafter for 4–5 weeks post-inoculation.

Indirect immunofluorescence microscopy was used to examine tick specimens after dissection when culture was not attempted. Antigen for IFA was prepared by growing B. burgdorferi stock cultures of strain ATCC 35210 / B31 grown in BSK-II medium in glass tubes in an incubator maintained at 35°C. Borrelia were harvested, centrifuged (International Equipment Co. Model CL, Needham Heights, MA, USA) at approximately 1500 rpm (352g) for 10 min and spirochaetes suspended above the sediment were removed with a sterile glass Pasteur pipette. An aliquot of the pellet was diluted 1 : 1 in PBS, centrifuged (Sorvall RC 5B Plus SS-34 rotor (Thermo Scientific, Walthan, MA, USA) at 10,000 rpm (7818g) for 10 min and the pellet resuspended in 1 ml PBS. Serial dilutions were prepared and a dilution selected that facilitated visualization of the spirochaetes without them overlapping in a glass well on Teflon-coated slides. The slides were air-dried and then frozen and stored at −20°C prior to use.

Suspect cultures and the gastrointestinal contents of dissected ticks were examined for B. burgdorferi by IFA (Levine et al., 1989) using monoclonal antibody H5332 (provided by Alan Barbour, University of Texas Health Science Center, San Antonio, TX, USA). Optimal working dilutions for each batch of conjugate were determined by draughtboard dilutions to accommodate for variations in the activity of each lot of antibody. Working dilutions of the monoclonal antibody ranged from 1/5 to 1/25. Working dilution for the fluorescein isothiocyanate conjugated anti-mouse immunoglobulin G antibody (Kirkegard and Perry Laboratories, Gaithersburg, MD, USA) was consistently 1/200 (Ouellette et al., 1997).

In 2005, ticks and tissues were ground with a sterile plastic pestle, resuspended in 100 µl of TE buffer and an IsoQuick Nucleic Acid Extraction Kit (ORCA Research, Inc., Bothwell, WA, USA) was used for DNA extraction (Levin et al., 2002). In 2009, ticks collected from three NC field sites were stored at −20°C until DNA extraction. Ticks were first individually washed with 70% ethanol. Extraction was carried out using a Qiagen DNeasy Blood and Tissue Kit (Qiagen, Valencia, CA, USA) using the modified, total DNA from ticks for the detection of Borrelia DNA protocol. The DNA samples were stored at −20°C until PCR testing was performed.

Genomic DNA extracted from the ticks collected and examined in 2009 was initially tested using tick-specific actin primers, to confirm that DNA was not degraded (Gao et al., 2011). Briefly, species-specific PCR primer sets were designed for B. burgdorferi, B. andersonii and B. bissettii with primerquest software (http://scitools.idtdna.com/Primerquest/) for amplification of the ospA gene (Table 1). Primers were purchased from IDT (http://www.idtdna.com). Polymerase chain reactions were run in 25 µl volumes with 12.5 µl AmpliTaq Gold^® Master Mix (Applied BioSystems, Foster City, CA, USA), forward and reverse primers at a final concentration of 250 nm,1 µl of template DNA and RT-PCR water (Ambion, Austin, TX, USA) to final volume. Amplifications were performed in an ABI GeneAmp 9700 (Applied BioSystems, Foster City, CA) thermal cycler with the following conditions: 95°C for 5 min; 40 cycles of 95°C for 60 s, 59°C for 60 s, 72°C for 120 s; 72°C for 7 min followed by a 4°C hold for the duplex reaction containing B. burgdorferi and B. andersonii primers. The uniplex PCR reaction contained B. bissettii primers only and was run as above except the annealing temperature was increased to 70°C. Polymerase chain reaction products were electrophoresed in 2% agarose gels stained with SYBR^® Safe (Invitrogen, Carlsbad, CA,USA) and visually recorded with a BioDoc-It (UVP, Upland, CA, USA) gel documentation unit. The PCR amplicons were excised from agarose gels, purified (Qiaquick Gel Extraction kit; Qiagen, Valencia, CA, USA) and sequenced (McLab, S. San Francisco, CA, USA). NCBI BLAST searches were performed with sequenced amplicons to verify identities of the genes.

Sixteen Borrelia isolates obtained in 2005 from ear punch biopsies and held at −80°C were thawed and placed in BSK-H culture medium as described above. Although other isolates had been obtained previously to 2005, they were no longer available for extraction and sequencing. Extraction of DNA from presumed Borrelia cultures was performed using DNeasy Blood and Tissue Kit (QIAGEN, Valencia, CA, USA) following the manufacturer’s protocol. A maximum of 2 × 10⁹ cells/ml were used for DNA extraction. Concentration and quality of the extracted DNA was determined with a Nanodrop 1000 (Thermo Fisher Scientific, Inc., Waltham, MA, USA). The extracted DNA was stored at −20°C prior to preparation for amplification.

To verify that each of the 16 Borrelia culture only contained a single genospecies, prokaryotic 16S rRNA gene amplicons were prepared following the 16S Metagenomic Sequencing Library Preparation protocol for the Illumina MiSeq system with some modifications. Briefly, primers were designed to amplify the V3 and V4 regions of 16S rRNA (Muyzer et al., 1993; Sim et al., 2012) and with overhang adapter sequences compatible with the Illumina index and sequencing adapters (Table 2). Polymerase chain reaction amplification components were NEBNext High-Fidelity 2× PCR Master Mix (New England Biolabs, Ipswich, MA, USA), 1 µm of each amplicon PCR primer and 5 ng/µl template DNA. The amplification PCR was performed using: one cycle of 95°C for 3 min; 25 cycles each of 95°C for 30 s, 55°C for 30 s and 72°C for 30 s; 1 cycle of 72°C for 5 min; and hold at 4°C. Clean-up of the PCR amplicon products to remove free primers and primer dimers was performed using Agencourt AMPure XP beads (Beckman Coulter, Brea, CA, USA). Following amplicon clean-up, second set of PCR was performed to attach barcoding indexes to the amplicon PCR products. Dual-index primers were designed so that samples could be multiplexed in one MiSeq lane. The index PCR was performed as follows: one cycle of 95°C for 3 min; eight cycles each of 95°C for 30 s, 55°C for 30 s, and 72°C for 30 s; one cycle of 72°C for 5 min; and hold at 4°C. Clean-up of the PCR index products was performed as described above using Agencourt AMPure XP beads. The indexed amplicon libraries were normalized and pooled. The pooled library was checked for quality and quantified using a High Sensitivity DNA chip on the Agilent 2100 Bio-analyzer (Agilent, Santa Clara, CA, USA). The library was diluted and combined with a PhiX Control library (v3) (Illumina, San Diego, CA, USA) at 10% and sequenced on two lanes on an Illumina v3 300PE MiSeq sequencer (Illumina), using standard sequencing protocols. Base calls were generated on instrument during the sequencing run using the MiSeq Real-Time Analysis (RTA 1.18.54) software. FastQ generation, demultiplexing and adapter trimming were performed by the MiSeq Reporter software (2.4 and 2.5.1).

The resulting demultiplexed reads were processed using the QIIME 1.8.0 toolkit (Caporaso et al., 2010). Briefly, the paired end reads were joined together and open reference-based OTU picking was performed where Uclust (Edgar, 2010) was applied to search using Greengenes 13_8 reference database (DeSantis et al., 2006) filtered at 97% identity. Reads not matching a reference sequence were removed from analysis. OTUs were assigned based on a database hit of 97% or greater sequence identity, and taxonomy was assigned against the appropriate database.

Genomic DNA from 65 randomly selected Ixodes ticks from Nags Head Woods Preserve (n = 25), Corolla (n = 21) and Pine Island (n = 19) collected in October 2009 were reexamined for Borrelia by amplifying 5S–23S intergenic spacer (IGS) region of Borrelia spp. using the specific primers 23SN1 and 23SC1 (Rijpkema et al., 1995). PCR reactions were each done in a 20 µl volume comprised of 10 µl of AmpliTaq Gold PCR Master Mix, 1 µl (10 µm) each of respective forward and reverse primers, 1 µl of template DNA and 7 µl of nuclease water. Thermocycler conditions for amplification of 5S–23S IGS fragments were as follows: 95°C for 10 min, 39 cycles of 94°C for 1.00 min, 50°C for 1.30 min, and 72°C for 1.30 min, and a final cycle of 72°C for 5 min. Genomic DNA from 16 Borrelia cultures isolated from ear punches of O. palustris and P. leucopus in 2005 were also amplified using the above-mentioned primers and PCR conditions.

Concurrently, species identifications of the field-collected I. scapularis adults were confirmed by amplifying mitochondrial 16S rRNA gene using the primers 16S+1 and 16S−1 (Black and Piesman, 1994). PCR reactions were prepared as previously described for the Borrelia 5S–23S intergenic spacer. The mitochondrial 16S rRNA gene of ticks was amplified using the following thermocycler conditions: Initial denaturation of 95°C for 10 min, 35 cycles of 95°C for 30 s, 54°C for 30 s, and 72°C for 1.00 min, and a final cycle of 72°C for 10 min. PCR products were electrophoresed in 1.2% agarose gels containing ethidium bromide and visualized through transillumination.

Amplicons from 16 Borrelia cultures from rodent ear punch biopsies taken in 2005 and four DNA samples from Ixodes collected in 2009 that produced expected sized bands of ~375 bp in Borrelia-specific 5S–23S PCR assay, and ~460 bp in 65 samples from tick mitochondrial 16S PCR assay were sequenced by Eton BioScience, Inc. (Research Triangle Park, NC, USA), using the forward primer that was used in amplification. Individual sequences from 5S–23S and mitochondrial 16S amplicons were analysed by BLASTN for sequence match analysis in NCBI database. The mitochondrial 16S rRNA gene sequences were multiple-aligned with ClustalX software package (Larkin et al., 2007) along with reference 16S rRNA gene sequences of I. scapularis strains retrieved from GenBank (2016). Haplotypes were identified by nucleotide sequence variation with each variant designated as a unique haplotype even if its sequence differed by only one substituted base (Norris et al., 1996).

A phylogenetic tree was constructed by the neighbour-joining method (Saitou and Nei, 1987) with bootstrap analyses consisting of 1000 iterations with MEGA 6 software package (Tamura et al., 2013), and evolutionary distances were calculated using Kimura 2 parameter model. All the sequences were submitted to GenBank under the accession numbers: Borrelia 23S: (ticks) KT821557–KT821560; (rodents) KX419243–KX419258; I. scapularis 16S: KT821561–KT821624.

Questing adult I. scapularis were collected at eight sites on the Outer Banks of North Carolina during 21 October 2011–March 1992, November–January 1993, November 1993, and October 2009 (Fig. 1). Sample collections traversed 134 km from Corolla, NC, to Cape Hatteras National seashore. Borrelia burgdorferi-infected I. scapularis were detected at four of eight (50%) sites in October 1991 (Table 3). Isolates were obtained from 34 questing adult I. scapularis collected from these four locations and confirmed to be B. burgdorferi by IFA with a species-specific monoclonal antibody (H5332). During February–March 1992, November 1992–January 1993, and November 1993, 541 questing ticks were collected at six locations on the Outer Banks, dissected and examined for spirochaetes by dark-field microscopy (Table 4). Seven percent of the questing adult I. scapularis were found infected, and the isolated spirochaetes were confirmed to be B. burgdorferi by IFA conducted with a species-specific monoclonal antibody (H5332). In October 2009, 71 adult I. scapularis were collected from three locations on the Outer Banks that had been previously sampled in the early 1990s (Table 5). Borrelia burgdorferi was detected by PCR (OspA) in 14% of the I. scapularis collected. The spirochaete was detected in two of the 24 (8%) adult questing I. scapularis collected in Corolla, and eight of the 23 (35%) adult questing I. scapularis collected on Pine Island. However, none of the 24 I. scapularis adults collected in Nags Head Woods were infected. All ticks tested were negative for B. andersonii and B. bissettii.

During 1991–1993, 304 rodents were live-trapped in 2546 trap-nights, and 58 rabbits (Sylvilagus floridanus, S. palustris) were live-trapped in 2301 trap-nights at Pine Island and Buxton Woods and examined for ticks. One hundred fifty-four six-lined racerunners lizards (Cnemidophorus sexlineatus) were also live-trapped in 13,573 coverboard checks, and 3,505 drift fence pit trap checks. Blood from 51 lizards, 58 rabbits and 45 rodents was inoculated into BSK-II medium. Spirochaetes were isolated from P. leucopus, O. palustris and S. palustris, but not C. sexlineatus trapped on Pine Island (Table 6). Isolates were obtained from 20% of the O. palustris and 50% of the P. leucopus, and 4% of S. palustris. During these studies, a small number of I. scapularis that survived removal from trapped hosts were examined for spirochaetes by IFA. No spirochaetes were observed in 8 larvae removed from P. leucopus trapped at Pine Island. However, spirochaetes were detected in one of two nymphs removed from P. leucopus, and one of three larvae and one of four nymphs removed from O. palustris trapped on Pine Island during October and November 1992. No spirochaetes were observed in one nymph and six larvae removed from P. leucopus or in a nymph removed from a south-eastern five lined skink (Eumeces inexpectatus) trapped at Buxton Woods.

In another study conducted during August 1993–September 1994, three species of small rodents (n = 123) trapped in 1491 trap-nights were examined for Borrelia. Borrelia burgdorferi was detected in 36.6% of the trapped rodents. The majority of rodents trapped were white-footed mice (n = 106) (Table 7), and with the exception of January 1993, Borrelia-infected P. leucopus were trapped throughout the 13 consecutive months in which trapping was conducted. Eighty-nine B. burgdorferi isolates were obtained from the various tissues collected and confirmed by IFA. Borrelia were predominately isolated from bladders and skin samples. No spirochaetes were found in the embryonated tissues; however, B. burgdorferi was isolated from one of the two pregnant white-footed mice from which these tissues were obtained. One of three (33.3%) Mus musculus a juvenile, trapped in April 1994 was infected. Fifty per cent of the O. palustris that were examined (n = 14) were infected with the spirochaete. Borrelia burgdorferi was detected in one adult male O. palustris; but not a juvenile male trapped in November 1993. Two adult male O. palustris (100%) were found infected in December 1993, but the spirochaete was not observed in two adult females. In addition, the spirochaete was detected in one adult male O. palustris that was one of four rice rats trapped (25%) in February 1994, an adult male rice rat trapped in April and three (100%) adult male rice rats trapped in August 1994. Borrelia burgdorferi was detected in 37 of 106 (35%) P. leucopus (Table 7). The spirochaete was detected in P. leucopus in all months except January and was most prevalent in the mice in June. Four engorged ticks I. scapularis nymphs and 25 I. scapularis larvae were collected from infected rodents, but the ticks were not examined for spirochaetes.

Rodents, P. leucopus and O. palustris, were trapped at Pine Island again in May and July 2002. Fifty-five animals were trapped in 799 trap-nights. Blood and bladder tissue from 32 P. leucopus, and 11 O. palustris was placed in BSK-H culture. Spirochaete isolates were obtained from 4 (9%) of the animals examined. Spirochaetes were isolated from the bladders of three of 15 (20%) P. leucopus trapped in May 2002, but not in 17 P. leucopus trapped in July. The three infected animals were adult males. Spirochaetes were also isolated from one of 11 (9%) O. palustris trapped in July 2002. The animal was an adult male.

During March–May 2005 at Pine Island, 140 rodents and two small rabbits were trapped. Attached partially engorged I. scapularis were also collected from 18 rodent hosts [O. palustris (n = 3), P. leucopus (n = 15)]. Borrelia burgdorferi was detected by PCR (OspA) in 10 (69%) of 15 nymphal I. scapularis and 1 (20%) of 5 larval I. scapularis removed from hosts. Borrelia burgdorferi was not detected in two larvae removed from an adult O. palustris in March, but was detected in one larval I. scapularis removed from an adult male O. palustris in April. Borrelia burgdorferi was detected in one nymph removed from each of two adult female P. leucopus, and a third nymph from an adult male P. leucopus collected in April. The remaining ticks were removed from animals trapped in May. One infected nymph was found on a juvenile female P. leucopus and three adult male P. leucopus. Two infected nymphs were found on an adult female P. leucopus. Ear punch biopsies were obtained at the time of capture for culture. At approximately 72 h after culture preparation, Borrelia were observed via dark-field microscopy in ear punch cultures from 10 (29%) of 35 O. palustris and 15 (14%) of 105 P. leucopus. No spirochaetes were detected in blood cultures from two S. palustris. Twenty-nine animals were recaptured; four animals were trapped three times. One juvenile male white-footed mouse yielded positive cultures for spirochaetes both times it was trapped, first in April and then 21 days later in May.

The OTU profiles of 17 of the 26 spirochaete isolates obtained from rodent ear punch biopsies in 2005 were investigated using UCLUST consisting of 383,704 quality filtered MiSeq 16S rDNA sequencing reads. Using a sequence similarity threshold of 97% resulted in identification of one dominant OTU containing 380,430 reads (99.1%) across all of the 17 samples, indicating that the samples contained only a single Borrelia sp. Subsequent, nucleotide BLAST results for the GenBank database indicated that the sequences (GenBank Accession numbers: KU896130, KU896131, KU896132, KU896133) had 99–100% identity to B. burgdorferi strain B31 (Accession No. CP009656.1 accessed 2 February 2015; e-value <0.001). Sequence analysis of 5S–23S IGS amplicons from 16 of these isolates revealed that the Borrelia were 98–99% identical to GenBank sequences of the B31, JD and M11p strains of B. burgdorferi (Accession Nos. CP009656, CP002312.1 and, KM269459.1, respectively).

Of the 65 I. scapularis collected in 2009 and tested for Borrelia, four ticks from Pine Island were positive. BLAST analysis of the 5S–23S sequences showed that the ticks were infected with B. burgdorferi sensu stricto that exhibited 99% sequence similarity to the JD1 and B31 strain of B. burgdorferi (Accession Nos. CP009656.1 and CP009656, respectively).

The mitochondrial 16S rRNA gene fragments were successfully amplified by PCR from all 65 tick DNA samples. The sequences were 99% similar to the GenBank sequence of I. scapularis (Fig. 2), confirming the taxonomic identification of ticks collected on the Outer Banks as I. scapularis. The reconstructed neighbour-joining tree further showed that these 65 sequences of the 16S rRNA gene and GenBank reference sequences formed two major clades (Fig. 2). The Outer Banks I. scapularis sequences clustered into 12 haplotypes with nine haplotypes (58 ticks) and three haplotypes (seven ticks) represented in the two clades with significant bootstrap values. A majority of the I. scapularis ticks were assigned to Clade I containing sequences from northern and midwestern tick populations. Additionally, 18 of 19 I. scapularis from the Pine Island collection site, including four ticks infected with B. burgdorferi sensu stricto, were placed in Clade I. Notably, 16S rRNA gene sequences varied by 16–19 nucleotides for ticks collected at the same site (e.g. Nags Head Woods: Clusters C and P) as well as different sites [e.g. Pine Island (Cluster B) versus Corolla (Cluster R)] on the Outer Banks.